Post-incision events induced by UV : regulation of incision and the role of post-incision factors in mammalian NER Overmeer, R.M.

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Post-incision events induced by UV : regulation of incision and the role of post-incision factors in mammalian NER

Overmeer, R.M.

Citation

Overmeer, R. M. (2010, September 29). Post-incision events induced by UV : regulation of incision and the role of post-incision factors in mammalian NER.

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GG-NER complex assembly and regulation

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Chapter 2: GG-NER complex assembly and regulation

GG-NER is triggered by helix distortions

As mentioned, recognition of lesions by GG-NER is achieved by probing for helix distortion rather than recognition of the aberrant chemical moiety of the lesion per se. This is supported by the fi ndings that the repair rate is proportional to the amount of helix distortion as emphasized by the higher repair rate of the 6-4PP and N-(deoxyguanosine-8-yl)-2-acetylaminofl uorene (AAF) when compared to CPD and N-(deoxyguanosine-8-yl)-aminofl uorene (AF), which have a much smaller amount of helix distortion when compared to 6-4PP or AAF (Lee et al., 2004; Szymkowski et al., 1993; van Oosterwijk et al., 1998; Gunz et al., 1996). The dependence of repair on helix distortion is further emphasized by the observation that modifi cations such as C4′ backbone modifi cations are only repaired when combined with a mismatch, which distorts the backbone (Hess et al., 1997b) and that the repair of benzo[a]pyrene diol epoxide (BPDE) induced adducts is dependent on the opposing base, correlating repair with the helix distortion caused by the displacement of both the lesion and the complemen- tary base (Hess et al., 1997a). In the case of GG-NER, the XPC-HR23B-centrin2 complex is essential for the initial recognition of lesions (Sugasawa et al., 1998) however, the UV-DDB complex greatly enhances repair of 6-4PP and is indispensable for the repair of less helix distorting lesions such as CPD (Fitch et al., 2003; Moser et al., 2005). A recent publication by Scrima et al. (Scrima et al., 2008) provides a structural basis for the mechanism of lesion detection by the UV-DDB complex: the UV-DDB component DDB2 binds lesions through an ‘induced fi t’ mechanism in which the suboptimal stacking of the lesion containing helix leads to the appropriate conformation and fl exibility to fi t into the WD40 domain of DDB2.

DDB2 then further kinks the DNA and inserts a hairpin into the DNA fl ipping the lesion out.

The structure of DDB2 binding to lesions is compatible with chromatin bound DNA as the backbone of DNA wrapped around a histone shows a similar conformation to that of lesion containing DNA when bound by DDB2. The structure of lesion recognition within chromatin bound DNA correlates best when the lesion is situated in the outward facing minor grove, coinciding with where lesions are preferentially formed (Scrima et al., 2008; Gale et al., 1987). Furthermore, the UV-DDB complex is an ubiquitin ligase consisting of DDB1, DDB2, RBX1 and CUL4A and is associated with the COP9 signalosome which inhibits the CUL4 ubiquitin ligase activity. Exposure of cells to UV irradiation leads to activation of the ubiquitin ligase activity of the UV-DDB complex by dissociation of the COP9 signalosome, presumably by binding of DDB2 to damaged chromatin (Luijsterburg et al., 2007;

Shiyanov et al., 1999; Groisman et al., 2003). This leads to the ubiquitination of XPC at sites of damage, increasing its binding affi nity for lesions and facilitating the handover from UV-DDB (Sugasawa et al., 2005). In addition the UV-DDB complex may also ubiquitinate other proteins such as the histones H2A, H3 and H4 which would destabilize the chromatin, increasing accessibility to XPC and other NER components (Guerrero-Santoro et al., 2008;

Wang et al., 2006). The crystal structure of the S. cerevisiae XPC/HR23B homologues, Rad4/

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Rad23 respectively, sheds light on not only the recognition of lesions but also the handover of the lesion from DDB2 to XPC in mammalian cells (Min and Pavletich, 2007; Scharer, 2008). Whereas DDB2 binds the damaged strand, Rad4 initially binds the undamaged strand prior to probing for the helix distortion. As such the UV-DDB complex could increase the accessibility of the lesion by ubiquitination of H3 and H4 which would enable the binding of XPC to the undamaged strand. Prior to dissociation, UV-DDB would then ubiquitinate XPC, increasing its affi nity for the lesion and handing the lesion over to XPC.

Bi-partite recognition and assembly of the pre-incision complex

Recognition of the helix distortion is not suffi cient to trigger NER as, one to three basepair mismatch mediated, helix distortions without damage are not processed by NER (Hess et al., 1997b). Lesions consisting of a bulky adduct opposite dCMP deletions have even been found to lead to a dominant negative effect. Such damages apparently recruit the NER complex without triggering incision, trapping the NER complex and thereby inhibiting repair of other adducts (Buterin et al., 2002). Taken together, it is apparent that damage recognition by NER is bipartite requiring initial recognition of helix distortion and subsequent validation of the lesion. Initially this verifi cation was attributed to XPA in conjunction with RPA (Replication Protein A) (Evans et al., 1997b; Sugasawa et al., 1998; Volker et al., 2001). However, it has become increasingly clear that their role in NER lies more in recruitment, correct orientation and stabilization of other NER factors (see below)

XPC bound to helix distortions recruits the TFIIH complex. TFIIH is a basal transcription factor and consists of 10 subunits, which participates in promoter opening and clearance through the XPB helicase. Furthermore TFIIH contains a cyclin activated kinase which is able to phosphorylate RNAPII activating its transcriptional elongation function. In addition to the XPB helicase TFIIH contains the helicase XPD and it is the XPD helicase which is thought to be responsible for damage verifi cation. After recruitment of TFIIH by XPC through direct protein interactions with XPB and/or p62 (Yokoi et al., 2000), the helicase XPD unwinds the DNA in an ATP dependent manner creating a bubble structure required for the recruitment of other pre-incision factors. Although the helicase activity of XPB is dispensable for NER, its ATPase activity is not. Together with structural data from an archeal XPB homologue this suggests that XPB opens the DNA in an ATP dependent manner and that the helix is subsequently unwound by the helicase activity of XPD (Coin et al., 2007;

Fan et al., 2006). Structural data of the TFIIH complex suggest that the complex would stall at a lesion (Chang and Kornberg, 2000; Schultz et al., 2000). The proposed stalling of the XPD helicase activity at the lesion is the fi rst direct contact of the NER complex with the lesion and therefore verifi es the presence of a lesion, preventing incisions at mismatches and non-B DNA conformations (such as left-handed Z-DNA and tetraplexes) not containing lesions. Unwinding by TFIIH enables the recruitment of the other core NER factors XPA, XPG and RPA. The recruitment of these factors independent of each other however, their binding is synergistic as they stabilize each others binding in vitro (Riedl et al., 2003; Reardon and

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Sancar, 2003). The XPA dependent recruitment of XPF-ERCC1 signals completion of the pre-incision complex.

XPA and the requirements for XPF and XPG mediated incision

Whether XPA was the only prerequisite for the recruitment of XPF-ERCC1 (Volker et al., 2001) was not entirely clear as the binding of XPA to forked templates is greatly enhanced in the presence of RPA (de Laat et al., 1998; Matsunaga et al., 1996). However we recently showed that XPA can recruit XPF-ERCC1 in the absence of RPA, confi rming the notion that XPF-ERCC1 recruitment is strictly dependent on XPA (Overmeer et al., 2010b). As mentioned earlier XPA has been implicated as a damage verifi cation factor (Evans et al., 1997b;

Sugasawa et al., 1998; Volker et al., 2001). However binding assays show that XPA preferentially binds unusually kinked DNA structures such as those proposed to be formed by stalling of TFIIH at lesions (Dip et al., 2004; Missura et al., 2001). Moreover XPA increases the selective binding of the ssDNA binding protein RPA to the undamaged strand (Hermanson- Miller and Turchi, 2002), indicating that its role is in recruiting XPF-ERCC1 and correct positioning of RPA to the undamaged strand. In turn RPA plays an essential role in positioning XPF-ERCC1 and XPG, binding XPF-ERCC1 on its 5’ side and XPG on its 3’ orientated side.

Completion of the pre-incision complex triggers the endonucleases XPF-ERCC1 and XPG, incising 15-25 nucleotides 5’ and 3-9 nucleotides 3’ of the lesion respectively. The endonucleases incise in a near synchronous fashion (Moggs et al., 1996), therefore it has been diffi cult to determine the exact order of incision. Moreover there have been publications supporting both XPG independent incision by XPF-ERCC1 in vitro (Matsunaga et al., 1995;

Moggs et al., 1996) and vice versa (Mu et al., 1996; Evans et al., 1997b; Evans et al., 1997a) making the order of incision a subject of controversy and debate. However two recent papers (Mocquet et al., 2008; Staresincic et al., 2009) have shed new light on this. Mocquet et al.

showed that XPG has a slightly slower dissociation kinetics compared to other pre-incision NER factors in vitro and Staresincic et al. showed that ERCC1-XPF is able to incise in the presence of a catalytically inactive XPG in vitro whereas the reverse was not true. Further- more they showed recruitment of post-incision factors and partial gap fi lling in XPG defi cient cells (XPCS1RO) complemented with a catalytically defective XPG, both processes that require a 3’primer template formed by incision. In contrast, cells complemented with a catalytically defi cient XPF showed no recruitment of post-incision factors or repair synthesis at sites of local damage. These results implicate that 5’ by XPF-ERCC1 incision can take place prior to XPG mediated 3’ incision. The requirement of a catalytically inactive XPG for XPF- ERCC1 mediated incision confi rms the observation that XPG, in addition to its enzymatic activity, has a structural role in stabilizing the NER complex (Mu et al., 1997).

Transition from pre- to post-incision complexes

After incision an approximately 30bp gap needs to be fi lled by repair synthesis and the re- sulting repair patch must be sealed by ligation involving numerous post-incision factors. In

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contrast to most pre-incision factors the post-incision factors are not unique for NER but are

“borrowed” from other processes such as replication and BER (Table 1). As a consequence, it is diffi cult to elucidate the exact role of post-incision factors in NER and therefore the mechanism of transition from pre- to post-incision complexes has remained elusive. However, recently Mocquet et al. suggested that XPG and RPA are essential in mediating the transition form pre- to post-incision complexes. This is partially based on the observation that XPG has been shown to interact with PCNA (Gary et al., 1997), a homotrimeric ring that acts as a platform for a large number of proteins also known as the “sliding clamp” (Kelman, 1997).

Table 1. Overview of the NER associated proteins ‘borrowed’ from other processes, their origin, function and their NER associated function.

Factors involved in the post-incision steps of NER

Protein Pathway outside NER Function outside NER Function within NER RFCp36 Replication Associate together to form the core

of the clamp-loading complex

Idem

RFCp37 Replication

RFCp38 Replication

RFCp40 Replication

RFCp140 Replication Associates with RFCp36-RFCp40 to form clamp-loader which specifi cally (un)loads PCNA at 3’ DNA overhangs

Loads PCNA to sites of incision

Recruitment of Polδ

PCNA Replication Homo-trimeric circular ‘sliding clamp’ which acts as a mobile platform for replication and repair

Idem

Polδ Replication Main leading-strand polymerase Required for effi cient gapfi lling

Polκ TLS Translesion synthesis polymerase Required for effi cient gapfi lling

Ligase 1 Replication Main nick sealing ligase during replication

Able to seal nick in replicating cells

Ligase 3 BER Associate to form ligase which

seals nicks during BER

Main nick sealing ligase and required in quiescent cells

XRCC1 BER

CAF1 Chromatin assembly Cooperate in replication dependent H3/H4 deposition

Involved in chromatin assembly after gap-fi lling

ASF1 Chromatin assembly

Rad18 Protein modifi cation E3 ubiquitin ligase for PCNA Ubiquitination PCNA to enable recruitment of Polκ Ubiquitin Protein modifi cation Modulate protein stability, activity

or association

The role of RPA is thought to be through its interaction with RFC (Yuzhakov et al., 1999), a hetero-pentamer that opens and loads PCNA at 3’ primer templates (Majka and Burgers, 2004). This, together with the observation that the simultaneous presence of RPA and XPG is required for the repair synthesis, suggests that XPG and RPA play a pivotal role in the transition from pre- to post-incision complexes (Mocquet et al., 2008). The respective roles of XPG and RPA, through RFC, were elucidated further by the observation that knockdown

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of RFCp140 by siRNA led to a repair defect yet allowed recruitment of PCNA to sites of damage (Ogi et al., 2010; Overmeer et al., 2010a). This implies that the recruitment of PCNA is facilitated by XPG and does not require RFC. However, similar to normal replication (Lee and Hurwitz, 1990; Tsurimoto and Stillman, 1991), subsequent repair synthesis requires loading of PCNA by RFC as evidenced by the repair defect. Although PCNA recruitment seems to be facilitated by XPG, this recruitment requires incision as no difference in PCNA recruitment was seen in XP-G cells when compared to XP-A or XP-F cells, which are all defi cient in incision yet XP-G is recruited to sites of damage in XP-A and XP-F cells (Data not shown).

Cut-patch vs. cut-patch-cut-patch

The generally accepted model for gap fi lling is the so called cut-patch model which assumes that both incisions take place prior to gapfi lling (i.e. cut and then patch) (Aboussekhra et al., 1995; Araujo et al., 2000; Moggs et al., 1996; Mu et al., 1995). However, Staresincic et al.

have shown repair replication in cells unable to perform XPG mediated incision. Interes- tingly, the UDS of these cells, expressing a catalytically defi cient XPG, was only half that of normal cells or cells complemented with wt XPG. Based on this observation they go as far as to propose a novel model of cut-patch-cut-patch. Whereas the lower UDS observed led them to propose that 5’ incision is followed by partial gapfi lling only fi lling approximately half of the gap (Staresincic et al., 2009), the observed UDS could also represent only half of the repair sites being able to perform this cut-patch-cut-patch mode of repair due to a feedback loop (discussed later). The cut-patch-cut-patch model proposes 5’ incision followed by partial gapfi lling and subsequent 3’ incision, enabling completion of gapfi lling.

Interestingly, in support of this model, displacement synthesis at sites of UV repair has been suggested before albeit in the presence of DNA polymerase inhibitors and, at the time, the authors presumed that polymerase β was responsible for the observed displacement synthesis (Mullenders et al., 1985; Smith and Okumoto, 1984). However, as incisions are considered to take place in a near simultaneous manner (Moggs et al., 1996) and the requirement of DNA synthesis step between incisions would signifi cantly retard the second incision by XPG this cut-patch-cut-patch model has yet to be proven. Moreover, the experiments were performed in an artifi cially perturbed in vitro system; the exact requirements for XPG mediated incision in vivo have yet to be determined. It may well be possible that structural changes induced by XPF-ERCC1 mediated incision are suffi cient to trigger incision by XPG; alternatively the recruitment of post-incision factors and not the actual synthesis could be suffi cient to trigger incision by XPG. This is underlined by our observation that XPG was free to dissociate from sites of repair and associate to newly induced sites of damage under conditions where repair synthesis is inhibited (Overmeer et al., 2010b). Moreover, Mocquet et al. using a reconstituted NER system, showed coupling between the release of XPF and XPG with arrival of RFC and PCNA. These events occur prior to DNA synthesis, and hence their results are more consistent with the ‘classic’ cut-patch model.

Stirring things up, a recent paper (Ogi et al., 2010) showed some surprising dependency

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of NER on multiple polymerases to complete NER. Ogi et al. showed that knocking down either polδ or polκ led to an epistatic decrease in UDS of 50%. In addition RFC1 was found to be required for the recruitment of polδ, an observation we confi rmed in another publication (Overmeer et al., 2010a). Similarly polκ recruitment was found to depend on XRCC1 and ubiquitination of PCNA. The importance of RFC1 and PCNA ubiquitination in polymerase recruitment was confi rmed by knockdown of RFC1 and Rad18 which led to a similar defect.

The remaining UDS was found to require polε, interestingly recruitment was found to be dependent on CHTF18, an alternative clamploading complex, yet CHTF18 had no effect on UDS, suggesting CHTF18 inhibits polδ recruitment by RFC1 when not able to recruit and stimulate polε activity (Ogi et al., 2009). In conclusion it appears that post-incision is not performed in a single manner but that there are 2 distinct pathways active, each responsible for approximately half the repair synthesis (Ogi et al., 2010). It seems likely that the choice for each repair mode depends on a combination of factors such as the levels of nucleotides, (signalling induced) post transcriptional modifi cations of the polymerases or accessory factors and the chromatin structure present, whether this is partially through a cut-patch-cut-patch mode remains to be seen.

Repair replication and chromatin restoration

Irrespective of the cut-patch vs. cut-patch-cut-patch discussion repair replication initiates by the recruitment of PCNA and RFC. Loaded by RFC, PCNA enables the recruitment of most other post-incision factors such as DNA polymerases, ligases and chromatin remodelling factors (Moser et al., 2007; Mocquet et al., 2008; Yuzhakov et al., 1999), respectively required for repair synthesis, subsequent sealing of the nick and restoration of chromatin structure. Multiple DNA polymerases have been implicated in NER, including polδ, polε and polκ (Araujo et al., 2000; Moser et al., 2007; Ogi and Lehmann, 2006). However, the nature of their respective contribution to NER in vivo has, until recently, remained unclear. We showed that the siRNA mediated knockdown of a single polymerase led to a 50% decrease in unscheduled DNA synthesis (UDS or repair synthesis) (Ogi et al., 2010). PCNA ubiquitination by Rad18 was required for recruitment of polκ and therefore NER; this was surprising as PCNA ubiquitination was thought to be specifi c for translesion synthesis (TLS) and therefore S-phase cells (Lehmann et al., 2007). Furthermore polδ and polκ were epistatic, whereas polε was found responsible for the remaining synthesis.

Based on in vitro studies it was long assumed that Ligase1 was the primary ligase involved in NER (Aboussekhra et al., 1995; Araujo et al., 2000) however, later it was shown that XRCC1-Ligase3 was the predominant ligase in non-dividing cells and that Ligase1 was only involved in non-dividing cells (Moser et al., 2007).

After gapfi lling and ligation the chromatin complex must be restored through chromatin assembly factors such as CAF-1 and or ASF-1. This is thought to enable deactivation of the checkpoint and allow cell cycle progression (Chen et al., 2008; Gaillard et al., 1996; Green and Almouzni, 2003; Kim and Haber, 2009; Schulz and Tyler, 2006; Zhu et al., 2009). The

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role of chromatin and its modifi cations in repair has been reviewed recently and shall therefore not be discussed further (Dinant et al., 2008).

Points of no return and regulation

As mentioned above, damage recognition by NER is a bipartite process. This leads to the build-up of pre-incision complexes at sites with an altered base stacking but without damage (Hess et al., 1997b). When stably assembled, such complexes will have a negative impact on cellular functions by virtue of unwanted signalling and transcription/replication interference and hence need to be removed; in contrast, if damage is present the complex will be committed to repair at some point of the assembly process. Until recently it was not known which stage of the NER process comprises a “point of no return”. Unwinding of the helix by XPB and XPD has been suggested to be the fi rst irreversible step of NER (Gillet and Scharer, 2006); the recent observation that verifi cation of the damage occurs during the unwinding would validate this hypothesis. However, from a mechanistic view, incision is an obvious ‘point of no return’ as this step creates a gap which has to be fi lled. In a recent publication (Overmeer et al., 2010b) we showed that the pre-incision complex assembled in NER defi cient XP-A and XP-F cells is indeed stable as the proteins cannot be competed away by additionally induced damages. This would seem to support the notion that unwinding is indeed the fi rst point of no return. We also showed that in the absence of RPA, all pre-incision

Pre-incision factors

P OH

P

Dual incision RPA remains bound

P

OH Formation of abortive pre-

incision complex lacking RPA

Post-incision complex formation

Gap-filling and ligation is completed RPA and post-incision factors are able to dissociate

RPA is free to participate in novel incision events Pre-incision proteins

dissociate and bind to other lesions Damage recognition

Pre-incision complex formation

Post-incision factors

RFC

Polį

XRCC1 Lig3 Polİ Polț

XPC HR23b UV-DDB

TFIIH

XPA

XPG XPF

ERCC1

RPA RPA

RPA

RPA PCNA

RPA

Figure 1. Model of NER regulation by RPA cycling between synthesis and incision.

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factors associate with sites of damage indicating that the helix is unwound. However, despite the helix being unwound in the absence of RPA, the association of pre-incision factors with sites of damage is dynamic as the proteins could be competed away by additional sites of damage. This suggests that, instead of helix unwinding being the point of no return, the arrival of RPA stabilizes the complex and therefore commits the complex to excise the DNA lesion. The point of no return being prior to incision is especially important, as uncoupling incisions from repair synthesis and ligation could lead to an accumulation of ssDNA gaps which would be highly recombinogenic and persistently activate the DNA damage response.

Further emphasizing the importance of regulating NER, abortive formation of incision complexes already committed to incision would lead to an accumulation of complexes on DNA which, would in turn, pose a problem for replication, as shown for unrepaired single strand breaks by PARP inhibitors (Noel et al., 2006) and similar to that posed by (stalled) transcription complexes (Gottipati et al., 2008; Hendriks et al., 2008). Classic studies already revealed that incision is coupled to synthesis and ligation, as inhibition of synthesis led to a saturation

Figure 2. Flow diagram of the DNA damage signalling cascade.

DNA damage

Mediators

Transducers

Effectors

Cell cycle checkpoints Apoptosis Repair Transcriptional response Repair intermediates Stalled replication/transcription

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of incisions at low doses (Mullenders et al., 1985; Smith and Okumoto, 1984). This level of regulation was further corroborated by the observation that lesion removal is perturbed when repair synthesis is inhibited (Moser et al., 2007). Regulation of NER is independent of ATR mediated signalling events in non-dividing cells (Auclair et al., 2008; Overmeer et al., 2010b;

Vrouwe et al., 2010). We recently showed that the regulation of NER in non-dividing cells is mediated through RPA (Overmeer et al., 2010b). This regulation is based on RPA being the only factor known required for both pre- and post-incision steps. Moreover, there is only a limited amount of RPA available for NER which was apparent from the level of nuclear RPA decreasing in a UV dose-dependent manner. We showed that RPA, when sequestered in post-incision complexes, is unable to associate with pre-incision complexes. As we also showed that pre-incision complexes without RPA are unstable, these pre-incision complexes are abortive and would therefore not stably accumulate on chromatin, thereby preventing potential problems during replication. Surprisingly, release of RPA is not only dependent on synthesis but also sealing of the gap by ligation as inhibition of ligation by novel inhibitors (Chen et al., 2008) led to similar results. These results point to a mechanism which couples pre-incision complex stability and incision to repair synthesis and ligation, preventing accumulation of stalled pre-incision complexes, ssDNA gaps or nicks (Figure 1). As mentioned chromatin assembly is required to completely restore the chromatin structure to the initial state and deactivate cell cycle checkpoints. However, knocking down an essential factor such as CAF1 does not effect lesion removal or repair synthesis, indicating that regulation of incision is independent of chromatin remodelling (Polo et al., 2006). Yet it remains interesting to fi nd out how perturbed chromatin assembly, such as knocking down or inhibiting an essential factor such as CAF1, would affect signalling and cell cycle progression.

Interestingly, despite ATR having no role in regulating NER in non-dividing cells, Au- clair et al. showed that NER required ATR in replicating cells (Auclair et al., 2008). This dependence on ATR in replicating cells is distinct from the regulation of incision through RPA.

The affi nity of RPA for post-incision complexes containing ssDNA is higher than for pre- incision complexes, as evidenced by competition experiments (Overmeer et al., 2010b). Alt- hough RPA is present at much higher levels in replicating cells than quiescent cells, RPA has a (10-20 fold) higher affi nity for stretches of ssDNA formed during replication than a short ssDNA formed during repair as the former allows multiple RPA molecules to bind adjacently (Kim et al., 1994; Kim and Wold, 1995). Therefore, RPA would accumulate at replication intermediates and would not be able to effi ciently associate with pre-incision complexes. In addition, it was observed that RPA is released from PML bodies after UV damage induction (Park et al., 2005) and that the release of RPA from these PML bodies after damage requires ATR in S-phase but not G₁ cells (Barr et al., 2003). Together these observations suggest that the dependence of NER on ATR in S-phase cells is due to the ATR dependent release of additional RPA from PML bodies, further emphasizing the important role of RPA in regulating NER. Although release of RPA from PML bodies in an ATR dependent manner would explain the observed S-phase specifi c regulation of NER, phosphorylation of RPA might be able to

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achieve similar results. For it is known that RPA is hyperphosphorylated in response to DNA damage by ATR, ATM and/or DNA-PK (Binz et al., 2004; Block et al., 2004) and that such hyperphosphorylation diminishes the ability of RPA to stimulate replication in vitro, yet has no effect on its repair function (Patrick et al., 2005). Moreover phosphorylated RPA failed to associate with replication foci, yet associated with γH2AX foci in vivo (Vassin et al., 2004).

This leads to the hypothesis that hyperphosphorylation of RPA modulates the activity of RPA from replication to repair. However, closer examination of the respective protein interactions shows that hyperphosphorylation had no effect on the RPA-XPA interaction and that the decrease in stimulation of replication activity by hyperphosphorylated RPA correlated with a decrease of RPA-DNA polymerase α interaction (Patrick et al., 2005). As RPA is known to stimulate the DNA-polymerase α activity this suggests that hyperphosphorylation of RPA regulates initiation events and would therefore prevent fi ring of alternative origins after damage induction (Wold, 1997). Although it remains important to stress that hyperphosphorylation could also have a role in regulation of NER by ATR during S-phase, such regulation is probably predominantly through the ATR and damage dependent release of RPA from PML bodies, whilst damage induced hyperphosphorylation mainly serves to regulate replication.

Dynamics of repair

Much of work on the dynamics of NER has been done using GFP(variant)-tagged proteins (Alekseev et al., 2008; Hoogstraten et al., 2008; Houtsmuller et al., 1999; Luijsterburg et al., 2007; Mone et al., 2004; Nishi et al., 2009; Otrin et al., 1997; Solimando et al., 2009; Zotter et al., 2006). However, although these studies effectively measured the kon, koff and residence time of various proteins, recent data suggests that such fi gures are not directly correlated with a proteins function persé (Overmeer et al., 2010b). For example, the residence time of XPA was around 4-6 minutes and it was assumed that this refl ected the amount of time required for its participation (Rademakers et al., 2003). However, it seems unlikely that proteins would remain associated in vivo for extended time without covalent bounds. Indeed, when competition experiments were performed under conditions where no gap-fi lling took place XPA was able to associate with new damages despite no incisions taking place (Overmeer et al., 2010b), indicating that under these conditions dissociation of XPA does not necessarily require incision. In seeming contrast, similar experiments in NER defective cell lines suggested that XPA was unable to associate with other damages. This difference in association is due to the presence of RPA stabilising the complex, which subsequently requires incision for dis- solution of the complex and association of the individual proteins at novel sites of damage (Overmeer et al., 2010b). XPA was not the only factor to show such apparent immobilisation, RFC also remained associated with sites of repair in the presence of synthesis inhibitors.

However, despite being unable to associate with novel sites of damage, the residence time increased less than 50% when compared to non inhibited conditions (Overmeer et al., 2010a).

It would therefore seem that dynamic measurements of proteins in such complexes should be interpreted with some caution. Depending on the reaction speed, dynamic measurements can

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either represent the time required for the protein to fulfi l its function or should be considered a measure of relative binding affi nity. The latter is especially true when the reaction speed is slow i.e. a large amount of time is required for the protein to fulfi l its function, as is the case for pre-incision factors in NER defi cient cells and both pre- and post-incision factors in the case of DNA synthesis inhibitors.

DNA damage induced signalling

To counteract DNA damage induced genetic alterations and cytotoxicity, the various pathways of DNA damage repair (Chapter1, Figure 1) are essential. Moreover, it is crucial that DNA damage is repaired before the cell progresses to replicate or undergoes mitosis to prevent mutagenesis and cytotoxicity. Hence additional regulation is required to enhance repair on one hand and to prevent progression of the cell cycle prior to completion of repair on the other.

Continuous monitoring of DNA damage and ongoing repair is essential to activate sig-

Figure 3. A simplifi ed model of the sequence of UV damage induced signalling activation.

ATR ATRIP RPA R R R RPAPAAA

9-1-19-1111--11-9

Rad17d 7 Ra Raadadd 7d17

Claspin TopBP1 ATR ATRIP

RPA R R R RPAPAAA

9-1-19-1111--11-9

ATR

RPA R RP R RPAPA

ATRIP RPA R RP R RPAPA

Rad17d 7 Ra Raaa 1add 7d1

Rad17d 7 Ra Raadadd 7d17

9-1-19-11-9-91-9-

Claspin TopBP1

+ Chk1 +

Chk1 ATR

XPA p53 Repair stimulation CDC25 Rad18 checkpoint activation

ATR ATRIP RPA R RP R RPAPA

?

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nalling pathways that communicate with and control cell cycle progression. However it was and is not fully understood how the signalling apparatus senses damage, as the types of DNA damage that activate signalling are diverse and no conclusive evidence for signalling specifi c factors recognizing damage (i.e. sensors) was found. As although some reports have suggested that ATR can bind damaged DNA directly in vitro (Unsal-Kacmaz et al., 2002; Choi et al., 2009), in vivo results contradict such direct recognition of damage (Vrouwe et al., 2010). In recent years it has become clear that the initiation of signalling depends largely on recognition of repair intermediates as opposed to the damages directly (Cimprich and Cortez, 2008) However, the effect of DNA damage on continuous cellular processes, such as stalling of transcription or replication can also initiate signalling (Rouse and Jackson, 2002; Fousteri and Mullenders, 2008). Therefore, the majority of signalling activation is dependent on the recognition of DNA damage by the repair pathways or its interference with ongoing cellular processes. ATM and ATR are the main transducers (complexes that have a central function in signalling) and can recognize some of these repair intermediates directly although mediators, such as TopBP1, can also recruit the transducers to sites of repair. Finally, the transducers directly or indirectly activate the effectors such as p53. The effectors in turn activate cell cycle checkpoints, induce/ inhibit transcription of specifi c genes, stimulate repair and, if necessary, activate the apoptosis pathway (programmed cell death) (Zhou and Elledge, 2000) (Figure 2).

The main transducers involved in damage signalling are the PIKKs (phosphoinositide 3-kinase related protein kinases) such as ATM (ataxia telangiectasia mutated), ATR (ataxia telangiectasia and Rad-3-related), and SMG1 (suppressor with morphological effect on geni- talia family member). Where ATM activation is generally triggered by double strand breaks (DSB), ATR is the main transducer activated by stalled replication and damages repaired by NER such as UV photolesions (Abraham, 2001). The method of activation of SMG1 is not yet understood. However it seems to be activated in a similar manner as ATM (Brumbaugh et al., 2004; Gehen et al., 2008).

ATR is recruited to sites of damage by interaction of ATRIP (ATR-interacting protein) with repair intermediates containing RPA coated ssDNA (MacDougall et al., 2007; Zou and Elledge, 2003). However, activation of ATR, and subsequently the downstream kinase Chk1, also requires the loading of the 9-1-1 complex (a PCNA-like sliding clamp consisting of Rad9-Rad1-Hus1) by the Rad17 complex (an RFC like clamploader where RFCp140 is re- placed by Rad17) (Majka et al., 2006a; Majka et al., 2006b) which in turn recruits TopBP1 and Claspin (TopBP1 activates ATR, whilst Claspin activates and recruits Chk1) (Kumagai et al., 2006; Lee et al., 2005)(Figure 3). Although Chk2 is mainly activated by ATM in the presence of DSB, there are indications that Chk2 may be activated independent of ATM (Li and Stern, 2005; McSherry and Mueller, 2004).

Once activated, ATR and the effector kinases (Chk1 and Chk2) can regulate the cell cycle through a plethora of factors, either directly by ATR itself, such as p53 and XPA (reviewed by Cimprich and Cortez (Cimprich and Cortez, 2008)), or by Chk1 and Chk2, such as the CDC25 family and Rad18 (reviewed by Stracker at al (Stracker et al., 2009)).

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Intriguingly a novel method of preventing UV induced mutagenesis in NER defi cient cells was recently described. In the absence of NER Ape1 incises at 6-4PPs, yet no repair synthesis is observed. The incision of Ape1 is usually followed by removal of the damage 5’

to 3’ by Fen1. However, Fen1 normally cleaves single bases and removal of a 6-4PP requires removal of 2 bases. As such the incision by Ape1 would lead to an un-repairable DNA structure containing a single strand break. Due to the single strand break this structure is able to induce signalling, thereby preventing replication and thereby mutagenesis(Vrouwe et al., 2010). This is supported by the Ape1 haplo-insuffi ciency observed in XPC defi cient mice (Cheo et al., 2000)

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